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Translation Stress Positively Regulates MscL-Dependent

Excretion of Cytoplasmic Proteins

Rosa Morra,

a

Francesco Del Carratore,

a

Howbeer Muhamadali,

a

Luminita Gabriela Horga,

a

Samantha Halliwell,

a

Royston Goodacre,

a

Rainer Breitling,

a

Neil Dixon

a

aManchester Institute of Biotechnology, School of Chemistry, University of Manchester, Manchester, United

Kingdom

ABSTRACT

The apparent mislocalization or excretion of cytoplasmic proteins is a

commonly observed phenomenon in both bacteria and eukaryotes. However,

re-ports on the mechanistic basis and the cellular function of this so-called

“nonclassi-cal protein secretion” are limited. Here we report that protein overexpression in

re-combinant cells and antibiotic-induced translation stress in wild-type

Escherichia coli

cells both lead to excretion of cytoplasmic protein (ECP). Condition-specific

metabo-lomic and proteomic analyses, combined with genetic knockouts, indicate a role for

both the large mechanosensitive channel (MscL) and the alternative ribosome rescue

factor A (ArfA) in ECP. Collectively, the findings indicate that MscL-dependent

pro-tein excretion is positively regulated in response to both osmotic stress and

arfA

-mediated translational stress.

IMPORTANCE

Protein translocation is an essential feature of cellular organisms.

Bacteria, like all single-cell organisms, interact with their environment by

transloca-tion of proteins across their cell membranes via dedicated secretransloca-tion pathways.

Pro-teins destined for secretion are directed toward the secretion pathways by the

pres-ence of specific signal peptides. This study demonstrates that under conditions of

both osmotic stress and translation stress,

E. coli

cells undergo an excretion

phe-nomenon whereby signal peptide-less proteins are translocated across both the

in-ner and outer cell membranes into the extracellular environment. Confirming the

presence of alternative translocation/excretion pathways and understanding their

function and regulation are thus important for fundamental microbiology and

bio-technology.

KEYWORDS

ArfA, MscL, osmotic stress, protein excretion, translation stress

B

acteria use a variety of secretion systems to translocate proteins across their cell

membrane(s). These systems are involved in a range of essential cellular functions

facilitating bacterial virulence, cell attachment, and nutrient scavenging. In

Gram-negative bacteria, six secretion pathways (T1SS [type 1 secretion system] to T6SS) have

been identified. The T1SS, T3SS, T4SS, and T6SS pathways all translocate proteins in a

single step from the cytoplasm to the extracellular environment. In contrast, the T2SS

and T5SS pathways secrete proteins in a two-step process in which proteins are first

translocated across the inner membrane (IM) by either the general secretion (SecYEG)

pathway or the twin arginine (Tat) pathway and then across the outer membrane (OM)

in a T2SS/T5SS-dependent manner (1). In addition, a number of periplasmic

Escherichia

coli

proteins have been discovered to be translocated to the extracellular medium

without being associated with one of the classical OM secretion pathways under

standard laboratory cultivation conditions (2, 3). A considerable body of evidence

suggests that bacterial cells are also capable of releasing proteins to the extracellular

environment in the absence of a signal sequence, and these observations are

collec-Received17 November 2017Accepted3 January 2018Published30 January 2018 CitationMorra R, Del Carratore F, Muhamadali H, Horga LG, Halliwell S, Goodacre R, Breitling R, Dixon N. 2018. Translation stress positively regulates MscL-dependent excretion of cytoplasmic proteins. mBio 9:e02118-17.

https://doi.org/10.1128/mBio.02118-17. EditorSang Yup Lee, Korea Advanced Institute of Science and Technology

Copyright© 2018 Morra et al. This is an open-access article distributed under the terms of theCreative Commons Attribution 4.0 International license.

Address correspondence to Neil Dixon, neil.dixon@manchester.ac.uk.

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tively referred to as nonclassical secretion (4, 5), excretion of cytoplasmic protein (6, 7),

or moonlighting proteins (8). Nonclassical secretion has primarily been observed in

Gram-positive bacteria; however, analysis of the extracellular proteome of laboratory

E. coli

strains has identified a significant number of cytoplasmic proteins (9). To date, no

generalized mechanism has been elucidated and the topic is controversial, in particular

as a number of cytoplasmic proteins characterized as being released via

nonclas-sical secretion have subsequently been found to be released by OM vesicles (OMVs)

(5, 10, 11).

In this study, we observed that during

E. coli

cell culture, signal peptide-less

recombinant protein is localized in the medium at a titer of 0.7 g/liter and up to 80%

purity in a lysis-independent manner. Analysis of growth medium osmolality and the

metabolic footprint indicates that a hypo-osmotic stress is linked to the excretion

phenomenon. We observed that proteins are excreted across the IM into the

periplas-mic space in a large-conductance mechanosensitive channel (MscL)-dependent

man-ner. Condition-specific proteome analysis additionally found translation stress response

signatures to be highly associated with this excretion, which was validated by genetic

knockout. Collectively, the findings suggest a direct linkage of osmotic stress, the

alternative ribosome rescue factor A (ArfA)-mediated response to translational stress,

and MscL-dependent excretion. Finally, the findings were also validated in a wild-type

(nonrecombinant) background, confirming the physiological relevance of the excretion

phenomenon.

RESULTS

Recombinant proteins observed in

E. coli

culture medium during late

expo-nential growth phase.

In the expression of signal peptide-less recombinant proteins

in an

E. coli

BL21(DE3) host, enhanced green fluorescent protein (eGFP; 26 kDa),

3-hydroxybutyryl-coenzyme A dehydrogenase (AcDKR; 30 kDa), pectate lyase (PL;

44 kDa), IgG short-chain antibody fragment scFv13R4 (28 kDa), and transcription

termination/antitermination protein NusA (58 kDa) all unexpectedly led to extracellular

accumulation 10 to 12 h after induction (Fig. 1A and B). Protein quantification showed

a yield of ~0.7 g/liter (90 mg/g of dry cell weight) and a relative purity of 60 to 80% at

20 to 24 h postinduction (see Fig. S1A in the supplemental material). Importantly, all of

the recombinant proteins tested retained their corresponding biological activity

(Fig. S1B and C) and localized in the extracellular medium with relatively high purity.

Extracellular localization of recombinant protein is not due to cell lysis.

General

cell lysis was assessed by examination of the protein profiles of different cellular

fractions at 20 h postinduction (Fig. 1C). The protein profile of the extracellular sample

is significantly different from that of total and soluble intracellular samples. This

suggests that lysis is not occurring, as this would result in the extracellular profile being

essentially identical to the soluble intracellular profile (Fig. S1D). In a CFU assay, no

significant difference in viability was observed between the uninduced and induced

conditions for the first 14 h postinduction, further indicating that the release of

recombinant protein is not due to cell death (Fig. 1D). In accordance with previous

reports, variation in viability among the lag, exponential, and stationary phases was

observed (12).

Extracellular localization of recombinant protein occurs from an existing

cel-lular pool.

To explore whether the lack of an observable effect on the cell growth

profile and cell viability are being masked by cell growth and lysis occurring at similar

rates, we performed cultivation in the presence of chloramphenicol (Cm). Cm was

added (8.5 h), after which cell density remained constant for 4 h, whereas slow growth

continued under the control condition (no Cm) (Fig. 1E, top). Under both conditions,

extracellular localization of eGFP was observed with no significant difference in the

detected signal (Fig. 1E, bottom). This was further confirmed by SDS-PAGE analysis of

the extracellular fraction where increasing eGFP protein is observed following Cm

addition (Fig. S1E). As no new cell growth or protein synthesis can occur in the presence

of Cm, this result indicates that the source of extracellularly localized recombinant

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protein is an existing pool of cells and already translated protein rather than

lysis-dependent release.

Cells undergoing extracellular release of recombinant protein display a

com-promised membrane phenotype.

To investigate cell membrane integrity during

cultivation, the permeability of the OM was assessed in a fluorescence-based

N

-phenyl-1-naphthylamine (NPN) uptake assay (13). Because of the incompatibility with eGFP, the

assay was performed with cells expressing AcDKR; a 6-fold increase in OM permeability

was observed in cells expressing the recombinant protein (Fig. 2A). The permeability of

the IM was assessed by monitoring the activity of the endogenous cytoplasmic

-galactosidase (

-Gal) in cells expressing eGFP (Fig. 2B). Minimal activity was observed

in the uninduced cells, whereas significantly greater activity (20-fold,

P

0.01) was

detected in induced cells expressing eGFP. Taken together, both the OM and IM results

indicate that the recombinant protein expression increases membrane permeability,

FIG 1 Analysis of extracellular protein profile and viability ofE. coliduring recombinant protein expression. (A) Extracellular protein profile ofE. coliBL21(DE3)pET-GOI during transition/early stationary phase. The recombinant protein is indicated by an asterisk. (B) Intracellular (Intra) and medium (M) localizations of eGFP and cell growth (OD600) over time monitored by SDS-PAGE (M) and RFU normalized to cell

density (RFU/OD600). (C) SDS-PAGE of the total intracellular fraction (T), the soluble intracellular fraction from sonicated cells (S), and the

extracellular fraction (E) at 20 h postinduction. (D) CFU assays in the absence (UI) and presence of induction (I, 250␮M IPTG) over time. (E) Cell growth (top) and extracellular eGFP localization (bottom) monitored after addition of Cm at the time indicated by the arrow. In panels B, D, and E, data are the mean of at least three biological replicates⫾the standard deviation.

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suggesting that this may be linked to extracellular localization of the recombinant

protein.

The membrane permeability of individual

E. coli

cells was further investigated by

measuring the uptake of the fluorescent dye propidium iodide (PI) by flow cytometry

(Fig. 2C). Although PI staining of cells is commonly referred to as a viability assay, a

number of studies of microbial hosts have shown that membranes are permeable to PI

under stressed conditions (14) and that PI-permeable membranes can be repaired (15).

Additional studies of

E. coli

have shown that cells become PI stained following

hypo-osmotic shock with no significant loss of viability (16). During the exponential and

transition growth phases (2 to 12 h), greater membrane permeability (up to 4-fold) was

observed in cells that were expressing recombinant protein (Fig. 2C), consistent with

the earlier OM and IM permeability analysis (Fig. 2A and B). To assess whether the

observed membrane permeability correlates with recombinant protein expression

levels, we imaged the colocalization of PI (staining) and eGFP (expression). The majority

of induced cells initially (2 h) demonstrated high eGFP expression accompanied by a

low level of PI staining (

75%) (Fig. 2D). The level of PI staining gradually increased

over time to a maximum at 12 h, at which point

50% of the cell population showed

both a high level of PI staining and a high eGFP signal level. In contrast, cultivation in

FIG 2 Permeability of the IM and OM ofE. coliBL21(DE3)pET-GOI during growth. (A) OM permeability (NPN uptake) analysis in the absence (black) and presence (gray) of AcDKR expression. (B) IM permeability (ONPG hydrolysis) analysis in the absence (dark) and presence (gray) of eGFP expression. (C) Cell permeability (PI uptake) analysis over time in the absence (black) and presence (gray) of eGFP expression. (D) FACS density plots of eGFP expression and PI uptake of cells expressing (GI, bottom) or not expressing (G UI, top) eGFP over time. UI and I, no addition and addition of 250␮M IPTG, respectively. Data represent mean values⫾the standard errors from independent biological experiments. a.u., arbitrary units.

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the absence of the inducer demonstrated a low level of PI staining and, as expected, a

minimal eGFP signal (Fig. 2D). In summary, these results indicate that extracellular

localization is linked to high expression levels rather than the cultivation conditions.

Signal peptide-less protein localizes in the periplasmic space before leading to

extracellular localization.

Next we explored whether the signal peptide-less

recom-binant protein(s) reaches the periplasmic space prior to being released into the

extracellular medium. Surprisingly, we consistently detected recombinant protein in

the periplasmic fraction during the exponential (Fig. 3A and B; Fig. S2A) and stationary

(Fig. S2B) phases. Recombinant overexpressed proteins NusA and scFv were also found

to localize in the periplasmic space (Fig. S2C and D). Potential cross contamination

during cellular fractionation was excluded by using antibodies against fraction-specific

markers (Fig. S2E).

E. coli

BL21(DE3) is a strain specifically engineered for recombinant

protein overexpression and carries deletions of the OmpT and Lon proteases and an

insertion of the highly active T7 bacteriophage RNA polymerase (RNAP) (17), which

confer enhanced expression capability (18, 19). To explore the strain dependency of the

phenomenon, we therefore also assessed the localization in

E. coli

K-12 strain BW25113

of recombinant protein expressed from an

E. coli

RNAP-dependent promoter (P

BAD

).

Recombinant protein was again detected in the periplasmic space of both NusA

(Fig. S6A)- and eGFP (Fig. 3C)-expressing strains, although the localization was greatly

reduced (14-fold) in comparison to that in the BL21(DE3) strain (Fig. S2B). In addition,

little or no protein was detected in the medium, indicating that the extent of

periplas-mic localization and extracellular localization is dependent on the expression level.

FIG 3 Analysis of recombinant protein localization. (A) Localization of eGFP in the periplasm (PP) over time in BL21(DE3) and representative Western blot analyses (insets). (B) Localization of eGFP in the spheroplast (SP) over time in BL21(DE3). (C) Localization of eGFP in the periplasm of parental, MscL-deficient (ΔmscL), and restored (ΔmscL/pmscL-wt)E. coliK-12 cells at 16 h postinduction and representative Western blot analysis (inset). (D) CFU assay of parental, MscL-deficient (ΔmscL), and restoredE. coliK-12 cells. eGFP fluorescence is shown as RFU normalized to cell density. Data are presented as the mean value⫾the standard deviation of at least three biological replicates.*,P⬍0.05;***,P⬍0.001;****,P⬍0.0001 (two-way ANOVA followed by Dunnett’s multiple-comparison test).

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Extracellular localization of recombinant protein is not linked to the formation

of OMVs.

It is known that part of the periplasmic space can be encapsulated inside

OMVs and its content can be found in the extracellular environment (20). Previous

studies have shown that OMVs can transport both periplasmic proteins such as the

-lactamase (21) and indeed abundant cytoplasmic proteins such as GroEL (11). We

therefore isolated OMVs as previously described (22) that showed a protein profile

similar to that previously reported (11), in particular, association with both OmpA and

GroEL (Fig. S2F). In contrast, eGFP found in the extracellular medium was not associated

with purified OMVs, indicating that the protein is not exported into the extracellular

environment via the action of OMVs.

Effect of culture medium on recombinant protein localization.

Since excretion of

protein into the periplasm was not observed in growth medium containing sodium

chloride (Luria broth [LB] or terrific broth [TB] supplemented with sodium chloride)

(Fig. S3A and B), we measured the osmolality of both TB and LB growth media before

and after cultivation. TB showed a lower osmolality (346 mOsm) than LB (461 mOsm)

preinoculation; further, the osmolality of the spent TB (16 h) decreased to ~250 mOsm.

In contrast, the osmolality of the spent LB was maintained (~450 mOsm) most likely

because of the nature and relatively high concentration of the sodium chloride

osmolyte (present at 170 mM). We thus sought to investigate if cellular stress caused

by the change in external osmolality in combination with the recombinant protein

expression were leading to the excretion phenomenon.

Metabolic profile and footprint indicate extracellular localization of

recombi-nant protein associated with lower levels of osmoprotectant.

The metabolic effects

of recombinant protein expression upon

E. coli

were assessed by using five different

induction conditions/strains by gas chromatography-mass spectrometry (GC-MS) of the

spent medium (metabolic footprinting) and cell extract (metabolic profiling) (see

Materials and Methods). The five specific strains/conditions assessed were an

E. coli

BL21(DE3) strain containing (i) an expression plasmid encoding the inducible green

fluorescent protein (eGFP) that has been induced leading to expression of the

recom-binant eGFP-encoding gene (GI), (ii) an expression plasmid encoding the inducible

transcription termination/antitermination protein NusA that has been induced (EI), (iii)

an expression plasmid encoding NusA that has not been induced (EU), (iv) no plasmid

(DE3,U), or (v) no plasmid in the presence of isopropyl-

-

D

-thiogalactopyranoside (IPTG)

(DE3,I). The BL21(DE3) strain containing eGFP, in the absence of induction, showed

significant expression (20% leak) and therefore was not used as a negative control.

Figures S4 and S5 highlight metabolites that were statistically significant and allowed

differentiation among the five different induction conditions/strains for both the

footprint and the profile (Fig. S3C and D). Most interestingly from the footprint,

putrescine (

P

7

10

⫺4

) was found to be negatively correlated with the induced

conditions (EI, GI) that result in extracellular localization of recombinant protein

(Fig. 4A). Putrescine is a polyamine and has been found to be associated with the

regulation of a number of bacterial stress response mechanisms, including oxidative

stress and the hyperosmotic shock response (23, 24). In the metabolic profile,

pu-trescine was found to correlate positively with plasmid presence (Fig. 4B). Taken

together, the lower external putrescine levels of the strains producing recombinant

protein (EI, GI) and the similar internal putrescine levels of the plasmid-containing

strains (EI, EU, GI) seem to indicate that the strains undergoing the excretion

phenom-enon are experiencing greater hypo-osmotic (less hyperosmotic) conditions.

Periplasmic localization requires large-mechanosensitive channel (MscL).

In the

literature, there are several reports of cytoplasmic proteins being released into the

periplasmic space upon growth medium exchange/osmotic downshock via the

mecha-nosensitive channel MscL (25, 26). However, other studies have suggested that protein

release is not dependent upon MscL but is an artifact of the fractionation procedure

(27). To explore the possible involvement of MscL in the observed excretion

phenom-enon, we compared the localization of the recombinant protein expressed from an

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E. coli

RNAP-dependent promoter (P

BAD

) in

E. coli

K-12 strain BW25113 (parental), an

MscL-deficient (Δ

mscL

) mutant strain (28), and a strain with MscL restored (Δ

mscL

mutant plus episomal

mscL

). The results show periplasmic localization of recombinant

protein occurring in the parental strain. In contrast, the Δ

mscL

mutant strain displayed

a significant decrease in periplasmic localization of eGFP, assessed by fluorescence

FIG 4 Significant differentially detected metabolites and proteins in cells undergoing excretion. (A and B) Metabolite analysis by GC-MS of putrescine from the medium (A) and cell extract (B) of parental (DE3), pET-NusA (E), and pET-eGFP (G) BL21(DE3) strains in the absence (UI) and presence (I) of IPTG. (C) Induction-specific proteins differentially detected by LC-MS analysis in eGFP-induced (GI) versus NusA-uninduced (EU) strains (red, upregulated; green, downregulated). (D) Induction-specific proteins differentially detected by LC-MS analysis in NusA-induced (EI) versus uninduced (EU) strains. (E) Comparison of proteins differentially detected under both conditions (C and D). In panels A and B, the data are mean values of five biological replicates⫾the standard deviation. In panels C to E, data are presented as fold changes [FC(log2)] in the normalized mean values for a minimum of four biological replicates of

proteins differentially detected with significance (P⬍0.05) testing performed by the Wilcoxon RS method.

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(5-fold;

P

9

10

⫺3

) (Fig. 3C), and a consistent decrease in NusA, assessed by Western

blot analysis, was also observed (Fig. S6A). Episomal expression of MscL rescued the

parental phenotype, and recombinant protein was again localized in the periplasmic

fraction (Fig. 3C). All of the deletion strains assessed demonstrated viability comparable

to that of the control strains and literature values (Fig. 3D) (29). We also created a

BL21(DE3)

mscL

deletion strain and again observed decreased (14-fold;

P

1.3

10

⫺3

)

periplasmic localization of eGFP in the absence of MscL (Fig. S6B). This result indicates

a direct role for MscL in the excretion of recombinant protein(s) into the periplasm of

E. coli

.

Proteome analysis.

Having confirmed that the underlying cause of the excretion

phenomenon is not cell lysis but that it is dependent upon MscL, we sought to explore

whether native regulatory mechanisms are involved; therefore, we investigated the

gene expression effects by performing an unlabeled proteomic study (see Materials and

Methods). We assessed the same three different expression conditions/strains (EU, EI,

GI) used for metabolomic studies, and for each condition, two separate sample fractions

(total cell extract and periplasm) were collected and analyzed to allow topological

information in addition to relative abundance to be assessed.

Protein identification in the cell extract.

Principal-component analysis (PCA) of all

15 protein profiles (three conditions, five replicates) was performed to characterize

condition clustering and identify any outliers (Fig. S7A and B). In total, 1,075 proteins

were commonly identified (with more than one unique peptide) from the cell extracts

under all of the experimental conditions tested (Table S1.1). Relative abundance and

differential expression profiles were identified following quantile normalization (QN)

(30) (Table S1.2). The differentially detected proteins were assessed with the Breitling

rank sum (RS) test (31) (Tables S1.3 to S1.5); for proteins significantly differently

detected under the different conditions (

P

0.05), along with descriptors and

anno-tations, see Tables S1.6 to S1.8. For the GI-EU comparison, 28 proteins were significantly

differentially detected (20 up, 8 down) (Fig. 4C); for the EI-EU comparison, 115 proteins

were significantly differentially detected (54 up, 61 down) (Fig. 4D). To identify

phenomenon-dependent effects and to remove effects due to the specific recombinant

protein (eGFP or NusA), differentially abundant proteins common to both the GI-EU and

EI-EU comparisons were selected (Fig. 4E). This identified 11 common proteins, 8 up, 2

down, and 1 oppositely detected (Table S1.9). For other comparison analyses of protein

abundances that are linked to the specific recombinant proteins, see Tables S1.10 and

S1.11.

GO and subcellular topology enrichment analysis of cell extract.

Enrichment

analysis was performed by the iterative group analysis (iGA) method (32) after the

annotation of detected proteins with gene ontology (GO) terms (AmiGO 2) (33).

Upregulated proteins in the GI-EU comparison displayed enrichment for proteins

annotated with ribosomal ontology terms (GO.0003735,

P

2.5

10

⫺10

; GO.0019843,

P

4.1

10

⫺9

) (Table S1.12). Similarly, upregulated proteins in the EI-EU comparison

displayed enrichment for proteins assigned to ribosomal ontology terms (GO.0003735,

P

1.3

10

⫺15

; GO.0006412,

P

4.9

10

⫺11

; GO.0019843,

P

6.5

10

⫺11

) and

downregulated proteins displayed enrichment for proteins of the OM-bounded

periplasmic space (GO.0030288,

P

3.2

10

⫺7

). The changes in protein abundance

were further explored by clustering the detected proteins into subcellular topological

clusters (34), and the relative total abundance for each topology cluster was plotted

(Fig. 5A). Upon induction (EI or GI), an increase in protein abundance associated with

ribosomal proteins (r) was observed, in addition to a decrease in periplasmic proteins

(G). These qualitative observations are further supported by iGA analysis of topological

enrichment (Table S1.13). Upregulated proteins in the GI-EU comparison displayed

enrichment for genes assigned with a ribosomal topology (r,

P

5.7

10

⫺11

), and

downregulated proteins displayed enrichment for a periplasmic topology (G,

P

2.9

10

⫺6

). Similarly, upregulated proteins in the EI-EU comparison displayed enrichment for

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proteins assigned with ribosomal topology (r,

P

1.5

10

⫺17

) and downregulated

proteins displayed enrichment for a periplasmic topology (G,

P

1.6

10

⫺13

).

Protein identification in the periplasmic fraction.

In total, 412 proteins were

commonly identified in the periplasmic fraction under all of the experimental

condi-tions tested (Table S2.1). For the significantly differently detected proteins (

P

0.05),

along with descriptors and annotations, see Tables S2.6 to S2.8. In the GI-EU

compar-ison, 15 proteins were significantly differentially detected (8 up, 7 down), and in the

EI-EU comparison, 20 proteins were significantly differentially detected (14 up, 6 down).

A

B

r

Ex EI

Ex EU

Ex GI

PP EU

PP EI

PP GI

G

r G

r G

G

G G

F1 F1

F1

C

S1 S2

S4 S5 S6 S7 S8 S9 S10 S11 S3

S12 S13 S14 S15 S16 S17 S18 S19 S20 S21 S22 S23 S24 S25 S26

GI/E U

EI/EUGI/EI ArfA-associated proteins

Translaonal Stress (Puromycin)

Translaonal Stress (Kasugamycin) 2.6e-09 4.3e-11 1e+00

1.9e-07 5.1e-10 1e+00

6.6e-06 7.3e-06 1e+00 GI/EU

GI/EU

GI/EU EI/EU

GI/EI EI/EU GI/EI

EI/EU

GI/EI

p value

Cyt

IM

PP

OM EC

D

FIG 5 Relative protein abundance with subcellular topological annotation and signature response heat map analysis. (A and B) Relative protein abundance for extract (A) and periplasm (B) clustered by subcellular topology in BL21(DE3)pET-NusA in the absence (EU) or presence (EI) of IPTG and in BL21(DE3)pET-eGFP in the presence of IPTG (GI). N, nucleoid associated; r, ribosomal; A, cytoplasmic; F1, peripherally associated with the IM facing the cytoplasm; B, integral IM proteins; F2, peripherally associated with the IM facing the periplasm; E, IM lipoproteins; G, periplasmic; I, OM lipoproteins; F3, peripherally associated with the OM facing the periplasm; H, integral OM proteins; F4, peripherally associated with the OM facing the extracellular space; X, extracellular space; un, unassigned; co, coannotated. (C) Enrichment analysis of upregulated proteome against stress response signatures S1 to S26 (Table S3) with expanded views of ArfA-associated proteins (S3) (36) and puromycin (S11) and kasugamycin (S18) treatments (37). (D) Periplasmic localization of eGFP in parental and ArfA-deficient (ΔarfA) BL21(DE3) confirmed by Western analysis (inset, representative of three biological replicates). Panels A and B, subcellular topology annotation (STEPdb) (34). Panel C, enrichment analysis performed by the iGA method.Pvalues were corrected for multiple testing by the Bonferroni method. In panel D, the data are the mean value⫾the standard deviation of at least three biological replicates.*,P⬍0.05;***,P⬍0.001;****,P⬍0.0001 (two-way ANOVA followed by Dunnett’s multiple-comparison test).

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As in the cell extract analysis, differentially abundant proteins common to both the

GI-EU and EI-EU comparisons (2 up, 2 down) were selected (Table S2.9). Interestingly,

among the cytoplasmic proteins that were found more abundantly in the periplasm,

DnaK, GroEL, and elongation factor Tu 1 (EF-Tu) have previously been described as

being nonclassical secreted proteins (5).

GO and subcellular topology enrichment analysis of the periplasmic fraction.

Enrichment analysis was performed as before (Table S2.12). Proteins less abundantly

detected in the GI-EU comparison displayed enrichment for proteins of the

OM-bounded periplasmic space (GO.0030288,

P

7.5

10

⫺13

). The more abundantly

detected proteins in the EI-EU comparison displayed enrichment of ribosomal proteins

(GO.0003735,

P

5.3

10

⫺8

), and less abundantly detected proteins were enriched for

proteins of the OM-bounded periplasmic space (GO.0030288,

P

8.6

10

⫺7

). The

changes in protein abundance/localization were explored further by clustering the

detected proteins into topological groupings (34), and the relative total abundance of

each topology cluster was plotted (Fig. 5B). An increase in the relative abundance of

proteins peripherally associated with the IM facing the cytoplasm (F1) under induced

conditions (EU, 12%; EI, 18%; GI, 31%) was observed. In addition, a decrease in the

relative abundance of periplasmic protein (G) from the uninduced conditions (EU, 62%)

to the induced conditions (EI, 42%; GI, 46%) was observed. Consistent with this

qualitative abundance analysis, iGA topology enrichment analysis demonstrated that

the less abundantly detected proteins in the GI-EU comparison displayed enrichment

for genes with a periplasmic topology (G,

P

1.0

10

⫺15

) (Table S2.13). For the EI-EU

comparison, more abundantly detected proteins displayed an enrichment of proteins

associated with the ribosome (r,

P

9.6

10

⫺9

), and less abundantly detected

proteins were associated with the periplasmic compartment (G,

P

2.0

10

⫺9

).

Signature analysis of the cell extract indicates that the excretion phenomenon

is associated with translation stress response.

To probe protein expression changes

further, an extensive number of enrichment analyses were performed against

previ-ously identified stress response signatures by using transcriptomic and proteomic

databases (35). By iGA enrichment analysis, a number of stress responses associated

with the observed changes in protein level were identified (Fig. 5C; Fig. S7C and

Table S3). The most significant enrichment (S3,

P

2.6

10

⫺9

) was observed between

proteins upregulated under induced conditions in our proteomics data and proteins

detected following pulldown with ArfA (Table S3) (36). Significant enrichments (S11,

P

1.9

10

⫺7

; S18,

P

6.6

10

⫺6

) were also observed among proteins upregulated

under induced conditions in our proteomics data that were enriched for proteins

encoded by transcripts that are upregulated following puromycin and kasugamycin

treatment, respectively (Table S3) (37). Both puromycin and kasugamycin are known to

target the ribosome and cause translation stress, and intriguingly, puromycin has also

been shown to alleviate growth arrest when SsrA is depleted in a Δ

arfA

mutant strain

(36).

Excretion is dependent upon ArfA under conditions of enhanced recombinant

protein expression.

As recombinant overexpression is greatly enhanced by T7 RNAP

(18, 19) and overproduction of recombinant proteins is known to cause translational

stress (38, 39), we explored the association between MscL-dependent excretion and

ArfA in the following strains: parental BL21(DE3) and ArfA deficient (Δ

arfA

mutant)

expressing eGFP (pET-eGFP), and parental K12 and ArfA deficient (Δ

arfA

mutant)

expressing eGFP (pBAD-eGFP). The results show a significant decrease in the

periplas-mic localization of eGFP in the BL21(DE3) Δ

arfA

mutant strain compared to that in the

BL21 (DE3) parental strain (3.2-fold,

P

5

10

⫺4

) (Fig. 5D), validating the association

of

arfA

with the excretion phenomenon. However, no significant difference in

periplas-mic localization was observed between the K-12 Δ

arfA

mutant strain and the K-12

parental strain (Fig. S6C), indicating that excretion is dependent upon

arfA

only under

conditions of enhanced recombinant expression. Finally as the

mscL

and

arfA

genes

overlap, Western blot analysis of the membrane fraction was performed to detect the

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presence of MscL in the BL21(DE3) parental and

arfA

-deficient strains, confirming the

association of the excretion phenomenon with the

arfA

gene (Fig. S7D).

Excretion of cytoplasmic protein is observed under conditions of translational

stress in wild-type cells.

To explore whether the observed excretion is an artifact of

recombinant protein expression or indeed a native physiological response, we

exam-ined if endogenous cytoplasmic proteins are similarly excreted from the cytoplasm of

wild-type cells under conditions of translation stress. Cm was used because it is known

to cause translation arrest (40) and is reported to cause upregulation of ArfA (41).

Medium osmolality was monitored, and Cm was added to cells during the late

exponential growth phase, when the medium osmolality had decreased (~Δ70 mOsm).

Following Cm addition, an MscL-dependent change in the periplasmic protein profile

was detected (Fig. S6D), consistent with the excretion observed in the recombinant

system (Fig. S2C). Specifically, Western blot analysis indicates that the cytoplasmic

protein EF-Tu accumulates in the periplasm over time upon the addition of Cm

(Fig. 6A). In agreement with previous reports (26), basal levels of EF-Tu were detected

in the periplasm in an MscL-dependent manner (Fig. 6B). However, upon the addition

of Cm to wild-type cells, a large increase in EF-Tu was observed in the periplasm

(16-fold,

P

5

10

⫺4

), and this excretion was also confirmed to be dependent upon

the presence of MscL. Attempts to induce translation stress with Cm in a Δ

arfA

mutant

strain led to failure of the cellular fractionation procedure, possibly indicating that

under conditions of translation stress, cells exhibit a compromised IM in the absence of

ArfA. Taken together, these results indicate that in both recombinant and wild-type

cells, the MscL-dependent excretion phenomenon is positively regulated under

condi-tions of translation stress via the action of

arfA

.

DISCUSSION

Here we report the discovery of a novel signal peptide-independent IM excretion

phenomenon in

E. coli

that is triggered by both osmotic stress and translation stress.

We have shown that abundant (recombinant) signal peptide-less proteins are released

into the extracellular medium in substantial quantities and that this is not associated

with a loss of cell viability. However, an increase in the permeability of both the OM and

IM was observed in cells exhibiting this phenomenon. Recombinant protein was

unexpectedly observed in the periplasmic space

3 h postinduction, prior to being

released into the extracellular medium in a manner not associated with OMV

encap-sulation. As previous studies have shown that during recombinant expression, the OM

FIG 6 Excretion of cytoplasmic protein EF-Tu in the periplasm following translation stress. (A) Western blot analysis of EF-Tu localization in the periplasmic (PP) fraction from parental E. coliK-12 in the presence of Cm (0.01 mg/ml) over time. (B) Periplasmic localization of EF-Tu shown as a percentage of the total (periplasm [PP] plus spheroplast [SP]) protein, in parental and MscL-deficientE. coliK-12 cells in the absence or presence of Cm (0.01 mg/ml) after 10 h of treatment. A representative Western blot analysis is shown as an inset. Data in panel B are presented as the mean value⫾the standard deviation of three biological replicates.***,P⬍0.001 (two-way ANOVA followed by Dunnett’s multiple-comparison test).

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can become compromised, releasing the periplasmic contents (42, 43), we focused here

on the IM phenomenon.

The observed phenomenon of recombinant protein excretion across the IM occurs

in an MscL-dependent manner when expression is driven by either a native

E. coli

RNAP

or bacteriophage T7 RNAP. MscL is known to protect cells against hypo-osmotic shock,

operating via a turgor pressure-dependent gating mechanism, and releases cellular

contents to avoid excessive cell expansion (44). The exact pore size of the MscL channel

is not known, and no X-ray structures of the open complex have been resolved to date.

However, the consensus pore size of ~30 Å, equivalent to a cross-sectional area of

~7 nm

2

(45), is consistent with the cross-sectional area of the proteins that we report

to be excreted across the IM in an MscL-dependent manner (ranging from 27 kDa/

~4.5 nm

2

[eGFP] to 58 kDa/~7 nm

2

[NusA]) (46). Because of the complexity of the

intracellular components, we were unable to measure the osmolality of the cell lysate.

However, differential detection of the hyperosmotic shock response osmolyte

pu-trescine suggests that the strains undergoing the excretion phenomenon are

experi-encing greater hypo-osmotic (less hyperosmotic) conditions. In addition, this notion is

consistent with the IM permeability observed in cells undergoing excretion (Fig. 2D)

and in agreement with previous studies showing that cells under conditions of

hypo-osmotic shock are stained by PI (16). During cell culture, we observed a decrease in

medium osmolality of ~100 mOsm. Even on the basis of presuming no change in the

internal osmolyte concentration, this would result in an increase in turgor pressure of

~2.5 atm (40 mOsm

~1 atm), above the gating threshold (~0.2 atm) observed for

MscL channels in isolated membranes (44). In contrast to previous studies, we have

found that medium exchange is not required to elicit MscL-dependent protein

excre-tion.

GO and subcellular topology analysis of proteins that increase in relative intracellular

abundance during the excretion phenomenon displayed a significant enrichment for

ribosomal proteins. This is surprising, as recombinant protein expression has previously

been shown to lead to depletion of ribosomal proteins (39, 47). Additionally, depletion

of amino acid is also known to lead to stringent-response-mediated depletion of

ribosomal proteins (48, 49). Proteins that decrease in abundance were found to be

significantly enriched with periplasmic proteins, consistent with the loss of the

periplas-mic contents during the observed phenomenon. Among the cytoplasperiplas-mic proteins

increasing in abundance in the periplasmic fraction, some were previously detected in

the extracellular space (8), including DnaK and GroEL (5).

The proteome signature of the excretion phenomenon was found to be significantly

associated with proteins that bind to ArfA (

P

2.6

10

⫺9

) and with the response to

antibiotic treatment targeting the ribosome (puromycin;

P

1.9

10

⫺7

). Interestingly,

puromycin is believed to act in a manner similar to that of ArfA in that they both enter

the A site of the ribosome and cause dissociation of the ribosome complex. Indeed,

puromycin and ArfA lead to premature nascent chain release and release of nonstop

mRNA from the ribosome, respectively (36, 50). Furthermore, puromycin has been

shown to alleviate growth suppression in an SsrA-depleted Δ

arfA

mutant strain (36). To

validate the association between

arfA

and the phenomenon, a genetic knockout of

arfA

was created in BL21(DE3), which led to significantly less excretion into the periplasm

(Fig. 5D). In contrast, when expression was driven from an

E. coli

RNAP-dependent

promoter (P

BAD

), no significant reduction of periplasmic excretion was observed in the

absence of

arfA

(Fig. S6C). This difference in the dependency upon

arfA

is most likely

due to the lower levels of recombinant expression, and hence reduced translation

stress, under control of the

E. coli

RNAP-dependent promoter. Therefore, in the absence

of

arfA

-mediated translation stress, MscL-dependent excretion is triggered by only

osmotic stress. Finally, and with the most relevance for native cell physiology, the

phenomenon was also observed in wild-type (nonrecombinant) cells undergoing

trans-lational stress (Fig. S6D). Specifically, the cytoplasmic protein EF-Tu was excreted into

the periplasm in an MscL-dependent manner (Fig. 6). Previously, basal levels of EF-Tu

have been detected in the periplasm following osmotic downshock (26). However, in

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this study, the basal excretion is greatly enhanced and is positively regulated in

response to translational stress in both recombinant and wild-type cells.

The findings presented here suggest that the observed MscL-dependent excretion

is a native response to translation stress via the action of

arfA

. Rescue of stalled

ribosomes is predominantly mediated by the

trans

-translation system through the

action of transfer messenger SsrA (tmRNA) and small protein B (SmpB). The

tmRNA-SmpB complex alleviates the stalling and targets the mRNA and nascent protein for

degradation (51, 52). When the

trans

-translation system is compromised, alternative

ribosome rescue systems A and B (ArfA and ArfB) are used. As the

trans

-translation

rescue pathway consists of a degradative function to recycle mRNA and amino acids,

whereas ArfA does not, it is tempting to speculate that MscL may possibly play an

excretory role in the ArfA-mediated response to translation stress. This would

presum-ably also result in the nonspecific release of cytoplasmic contents. Why cells would

utilize such an energetically inefficient rescue mechanism is not entirely clear.

Intrigu-ingly,

arfA

is located adjacent to

mscL

in the

E. coli

genome, an arrangement that is also

evolutionarily conserved across other bacterial species (53). Small RNAs from the

overlapping transcripts of both

arfA

and

mscL

have been detected (54, 55); detailed

mechanistic analysis will be required to dissect how posttranscriptional control may

regulate MscL-dependent excretion in response to osmotic and translation stress.

MATERIALS AND METHODS

Strains and plasmids used in this study.E. coliBL21(DE3) (Novagen) and K-12 BW25113 parent and Keio Knockout Collection strains (28), BL21(DE3) ΔarfAand ΔmscLmutants (generated in this work), pBAD (Thermo Fisher), pET expression vectors (Novagen), pRL128 (Addgene 40180), and pSIM18 (gift from SynBioChem, University of Manchester) were used in this study. The eGFP-encoding gene bearing a 5= His tag sequence was cloned into pET44 and pBAD, affording the pET44_eGFP and pBAD_eGFP vectors, respectively, in which no signal peptide or Nus tag was present. ThenusAgene bearing 5=and 3=His tags is present in the pET44 vector and was subcloned into the pBAD vector. The AcDKR-encoding gene (pET26b-AcDKR vector) was a kind gift from Reddy’s Laboratories (EU) Ltd. The pectate lyase gene (pET24a-PL) was a kind gift from Biocatalysts Ltd. The scFv13R4-encoding gene was synthesized (GeneArt) and subcloned into both pET44 and pBAD vectors.

Expression induction procedure.All cultures were grown in TB (2.7% yeast extract, 4.5% glycerol, 1.3% Bacto tryptone) or, where specified, in LB (10 g/liter NaCl, 5 g/liter yeast extract, 10 g/liter tryptone) supplemented with 0.2% glucose and inoculated directly from freshly plated single colonies. For exponential/transition growth phase experiments (0 to 12 h postinduction), precultures were grown at 20°C with shaking (180 rpm) to an optical density at 600 nm (OD600) of ~0.8 and then transferred to Ultra

Yield (UY) flasks (Thomson) with or without an appropriate inducer (I or UI). Cultures were then grown at 30°C with shaking (210 rpm) for up to 12 h. For late exponential/transition growth phase experiments (12 to 24 h postinduction), precultures were grown at 37°C with shaking (180 rpm) to an OD600of ~0.8

and then transferred to UY flasks with or without an appropriate inducer (I or UI). Cultures were then grown at 30°C with shaking (210 rpm) for up to 24 h. For the nonrecombinant system experiments, preinoculums were diluted to an OD600of ~0.01, transferred to UY flasks, and grown at 30°C with shaking

(210 rpm) for 14 h (OD600of ~20). Cm (0.01 mg/ml, final concentration) was added, and cells were

assessed at the time points indicated. The inducers 250␮M IPTG (Sigma), 13.3 mM arabinose (Sigma), and 1 mM rhamnose (Sigma) were used as required. The antibiotics ampicillin (100␮g/ml), kanamycin (50␮g/ml), and Cm (100␮g/ml) were used as required. A BMG ClarioStar microplate reader was used to measure the colorimetric fluorescence and cell density (OD600) of intact cells.

Expression analysis and quantification.To measure extracellular eGFP expression, cell cultures were harvested by centrifugation (4,500⫻g) and the relative florescence units (RFU) were assessed directly. To measure intracellular eGFP expression, harvested cells were washed twice in equal volumes of phosphate-buffered saline (PBS) with 0.2% Tween. RFU and OD600were measured, and normalized

(RFU/OD) values were plotted against the time postinduction. For SDS-PAGE and Western blot analyses, OD-normalized volumes were collected and resuspended in SDS-PAGE loading buffer. Equal volumes were loaded onto a 4 to 20% gradient gel (Bio-Rad) and separated by SDS-PAGE, and target proteins were confirmed by Western blot analysis. Membranes were blocked with 5% skimmed milk in PBS containing 0.2% Tween. His-tagged recombinant proteins (eGFP, scFv, NusA) were detected with a mouse anti-His monoclonal antibody (Thermo Fisher MA1-21315, 1:3,000 in 5% skimmed milk) and a IRDye anti-mouse IgG secondary antibody (LI-COR, 1:10,000 in 5% skimmed milk). Endogenous MscL protein was detected on total membrane (collected by ultracentrifugation from sonicated cells after growth in LB supplemented with 0.5 M NaCl) with an anti-MscL antibody (kindly provided by P. Blount, University of Texas Southwestern Medical Center). When required, anti-RNAP 70 (Abcam, Inc., ab12088, 1:3,000 in 5% skimmed milk) and anti-beta lactamase (Novus Biologicals, 8A5-A10, 10 ␮g/ml in 5% skimmed milk) were used as fractionation control. The infrared fluorescence of IRDye secondary antibodies was detected with an Odyssey imager (LI-COR). SDS-PAGE gels stained with Instant Blue (Expedeon) or Sypro red (Thermo Fisher) protein gel stain were assessed by densitometry analysis with

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ImageJ or Image Studio (LI-COR) software. Protein quantification was performed with purified recombi-nant protein and/or bovine serum albumin (BSA) standard curves. All data were measured in biological triplicates.

scFv-Gal binding assay.Two-microliter␤-Gal (0.3 mg/ml) dots were spotted directly onto a nitrocellulose membrane (Amersham Hybond). As a negative control, the same amounts of BSA were spotted onto the same membrane. The membrane was left to dry for 5 min. Nonspecific binding sites were blocked by incubating the membrane with 5% milk in PBS for 1 h (50 rpm, room temperature [RT]). The membrane was washed three times for 15 min with PBS and then left to dry. On each dot, the substrate was added as 2 ␮l of a serial dilution of the periplasmic fraction containing scFv13R4 (previously quantitated by Western blot analysis) and allowed to dry for 5 min. The membrane was blocked with 5% milk in PBS for 20 min (50 rpm, RT). His-tagged scFv13R4 protein was detected as described above for expression analysis and quantification. The signal intensity was quantified with the Image Studio 5.0 software for densitometry analysis and used for curve fitting by using a four-parameter logistic function. All data were measured in biological triplicates.

AcDKR activity assay.The extracellular fraction of a BL21(DE3)pET-AcDKR culture was used to assay the activity of the recombinant protein by a UV kinetic activity assay measuring the absorbance change at 340 nm because of the oxidation of NADH over a 5-min period. Ethyl 4-chloroacetoacetate (Sigma-Aldrich 180769) was used as the substrate. The activity of the enzyme was calculated in units per milliliter (micromoles per milliliter per minute) of NADH consumed. All data were measured in biological duplicates.

CFU assay. Cultures were grown as described above for the expression induction procedure. Ten-microliter drops of serially diluted (1:10) cultures were spotted onto agar plates containing the appropriate antibiotic. Colony formation was observed after incubation overnight at 37°C. The colonies were counted and normalized for an OD600of 1.0 to calculate the normalized number of CFU per

milliliter. All data were measured in biological triplicates.

OMV purification.OMVs were isolated as previously described (22). In brief, the following steps were performed. (i) The extracellular fraction was collected from UY cultures by low-speed centrifugation (4,500⫻gfor 15 min), which removed most of the bacteria, the rest of which were eliminated by sterile filtration through a 0.45-␮m filter. (ii) Centrifugation at high speed (35,000 ⫻ g) of the filtered extracellular fraction was followed by ultracentrifugation (200,000⫻g). (iii) The pellets (OMVs) were resuspended in 1 ml of 20 mM Tris (pH 8). Target proteins were confirmed by Western blot analysis with anti-OmpA (Antibody Research 111120, 1:20,000 in 5% skimmed milk), anti-GroEL (Sigma G6532, 1:20,000 in 5% skimmed milk), and anti-His (Thermo Fisher MA1-21315, 1:3,000 in 5% skimmed milk) antibodies. OMVs were prepared from biological triplicates.

IM permeability assays. E. coli BL21(DE3)pET44-eGFP was grown as described above for the expression induction procedure. At the postinduction time points indicated, ~109cells (OD

600of 1) were

collected, washed, and resuspended in 1 ml of PBS (~109cells/ml). Ten microliters (~107cells) were added

to 96-well microtiter plates containing 90␮l of 0.4 mg/mlo-nitrophenyl-␤-D-galactopyranoside (ONPG) substrate. The assay mixture was incubated at 30°C for 90 min (the reaction was stopped by adding 5 mM Na2CO3), ando-nitrophenol generation was monitored at 420 nm. Cells permeabilized with chloroform

(100␮l) and 0.1% SDS (50␮l) and incubated for 5 min at 28°C were used as a positive control. The data are expressed in blanked corrected absorbance units. All data were measured in biological triplicates. For fluorescence-activated cell sorting (FACS), cells were washed once and then diluted 1/1,000 in PBS. The fluorescent dye PI was added to the cells at a concentration of 7.4␮M. Flow cytometry was performed with a Sony SH800 Cell Sorter (Sony Biotechnology) equipped with a 488-nm laser, enabling excitation of eGFP and PI, with subsequent emission being measured through use of the 525- and 617-nm channels, respectively. Data were recorded for 100,000 cells per sample atⱕ5,000 events/s, fluorescence com-pensation was applied, and analysis was performed with SH800 software. Histograms of events versus eGFP and PI fluorescence were plotted and used to determine the mean fluorescence and coefficients of variation of the total population and gated subpopulations consisting of nonfluorescent and fluores-cent cells. Additionally, the perfluores-centage of total events falling within each gated subpopulation was recorded. All data were measured in biological duplicates.

OM permeability assays.E. coliBL21(DE3)pET26b-AcDKR was grown as described above for the expression induction procedure. At the postinduction time points indicated, ~109cells (OD

600of 1) were

collected, washed, and resuspended in 1 ml of PBS (~109cells/ml). Ten microliters (~107cells) was added

to 96-well microliter plates containing 90␮l of 20␮M NPN in PBS. The change in fluorescence at 420 nm following 30 min of incubation at 30°C was recorded. Permeabilized cells (OD600of ~1) with the addition

of 1 mM of EDTA and incubation at RT for 3 min were used as a positive control. The data are expressed in blanked corrected fluorescence units. All data were measured in biological duplicates.

Fractionation of E. coli. Cultures were grown as described above for the expression induction procedure. At the postinduction time points indicated, ~109cells (OD

600of 1) were collected, washed in

PBS, and resuspended in 250␮l of buffer I (100 mM Tris-acetate [pH 8.2], 500 mM sucrose, 5 mM EDTA). A 2.7-␮l volume of 15 mg/ml lysozyme (Sigma) was added, immediately followed by 250␮l of pure water (Milli-Q). The samples were incubated on ice for 5 min, after which 10␮l of 1 M MgSO4was added. The

spheroplast (pellet) and periplasm (supernatant) fractions were collected by centrifugation at 17,000⫻ gfor 10 min at 4°C. The periplasmic fraction was centrifuged again to remove any remaining sphero-plasts. The spheroplast fraction was washed twice in buffer II (50 mM Tris-acetate [pH 8.2], 250 mM sucrose, 2.5 mM EDTA, 5 mM MgSO4). All fractions were prepared from biological triplicates.

Knockout of ArfA and MscL in BL21(DE3).The knockout strains were generated as described previously for Lambda Red recombineering (56), with the pRL128 (template) (57) and pSIM18

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bination) (58) plasmids. The oligonucleotides used to amplify FRT-flanked kanamycin selection cassettes were H1_arfA(ACTGGCAGGATTACACTCGCGCCGTTAAATAACCAACTGGAGTTTTTATGTTGGGATCCGTCGA CCTGCA), H2_arfA(GATTTGCTGAAAGAGCAGAATAACCGCTCTTAACAAGCGCCTGAAAGCAGGAGGTGTAGG CTGGAGCT), H1_mscL(TTAACATTTGTTAGACTTATGGTTGTCGGCTTCATAGGGAGAATAACATGTTGGGATCC GTCGACCTGCA), and H2_mscL (ACCACTGGTCTTCTGCTTTCAGGCGCTTGTTAAGAGCGGTTATTCTGCTCGAG GTGTAGGCTGGAGCTGC).

Sample preparation for GC-MS.Samples for metabolic analyses (in five biological replicates) were taken from BL21(DE3) shake flask cultivations carrying either pET44_eGFP or pET44_NusA and grown for 16 h as described above for the expression induction procedure. Samples were collected for metabolic profile analysis by quenching 2-ml aliquots with a chilled 60% aqueous methanol solution (⫺48°C), followed by 10 min of centrifugation at 6,000⫻gat⫺9°C to harvest the biomass. Internal metabolite extraction was carried out with an 80% aqueous methanol solution (⫺48°C) and a freeze-thaw method as described previously (59, 60). A two-step derivatization protocol was employed for all samples, starting with oximation (with methoxyamine-hydrochloride in pyridine), followed by a silylation step with N-methyl-N-(trimethylsilyl) trifluoroacetamide (61, 62). For the footprint analysis, samples were collected by centrifuging 1 ml of medium at 6,000⫻gfor 10 min to remove the cellular biomass. The supernatant was collected, quenched in liquid nitrogen, and stored at⫺80°C. Upon analysis, 200-␮l aliquots of samples were transferred to new microcentrifuge tubes; this was followed by combining 100␮l from each of the samples to generate quality control (QC) samples (61, 63). A 100-␮l volume of an internal standard solution (0.2 mg/ml each succinic-d4acid, glycine-d5, and lysine-d4) was added to all samples

before they were dried overnight with a speed vacuum concentrator (Concentrator 5301; Eppendorf, Cambridge, United Kingdom).

GC-MS instrument setup and data processing.An Agilent 6890N GC oven (Agilent Technologies UK, Wokingham, United Kingdom) coupled to a Leco Pegasus III mass spectrometer (Leco, St. Joseph, MI) was used in conjunction with a Gerstel MPS-2 autosampler (Gerstel, Baltimore, MD) as previously described (63, 64). All collected data were deconvoluted with Leco ChromaTOF software, and the metabolites detected were identified in accordance with the Metabolomics Standards Initiative guide-lines (65). The chromatographic peak corresponding to IPTG was removed from the data before any further analysis to avoid any variation resulting from the presence of this compound. By using the QC samples, mass spectral features with high deviation and missing values were removed, followed by normalization of the peak areas according to the internal standard and OD600of each sample. All data

collected were analyzed with Matlab version 8 (MathWorks Inc., Natick, MA). All preprocessed GC-MS peak areas were initially subjected to PCA (66), followed by discriminant function analysis (PC-DFA) where required (67). PCA and PC-DFA loading plots were used to determine the main metabolites contributing to the clustering patterns. The statistical significance (P⬍0.05) of these metabolites was further confirmed by one-way analysis of variance (ANOVA) and Tukey’spost hoctest.

Sample preparation for proteomics.Samples for proteomic analyses (in five biological replicates) were taken from BL21(DE3) shake flask cultivations carrying either pET44-eGFP or pET44-NusA and grown for 16 h as described above for the expression induction procedure. Total cell extract (Ex) and periplasmic (P) samples were prepared as follows. (i) Ex samples were prepared from cells harvested at 4,500⫻g, washed in PBS (twice), resuspended in buffer 1 (10 mM Tris [pH 8] and 200 mM NaCl containing protease inhibitor cocktail and DNase), disrupted by sonication at 30 A for 10 min (20-s on, 40-s off cycle), and centrifuged at 35,000 ⫻ gfor 30 min. (ii) P samples were prepared as described above for the fractionation of E. coli. The samples were then incubated with GFP-Trap_A beads (ChromoTek) in accordance with the manufacturer’s protocol to deplete the dominant eGFP. SDS at 0.1% was added to the unbound fraction, which was snap-frozen in liquid nitrogen and stored at⫺80°C until further processing.

LC-MS/MS analysis. Samples were prepared for MS by a modified FASP method (68) with the following modification. Samples were buffer exchanged with 0.1% SDS in Microcon 30-kDa centrifugal filter units (Merck Millipore) at 14,000⫻g. After three cycles of buffer exchange with 0.1% SDS (50␮l), the protein concentration was measured with a Millipore Direct Detect spectrometer at AM3. A total of 12.5␮g of protein was added to a fresh 30-kDa filter tube for reduction (dithiothreitol), alkylation (iodoacetamide), and proteolytic digestion to be performed in the filter tubes. The SDS buffer in the samples was exchanged with 8 M urea– 0.1 M Tris-HCl buffer (pH 8) for two cycles. Following reduction and alkylation (68), the proteins were concentrated and resuspended in 6 M urea in Tris-HCl buffer (pH 8). Endoproteinase Lys-C was used for the initial digestion at a 1:30 enzyme-to-protein ratio; after 3 h, the urea concentration was dropped to 1 M with Tris-HCl (pH 8). The proteins were then digested with trypsin at a 1:50 enzyme-to-protein ratio. After digestion, peptides were collected by centrifugation and desalted with OLIGO R3 reversed-phase resin on a microplate system and then reconstituted in 5% acetonitrile and 0.1% formic acid. Digested samples were analyzed by liquid chromatography-tandem MS (LC-MS/MS) with an UltiMate 3000 rapid separation liquid chromatograph (RSLC; Dionex Corporation, Sunnyvale, CA) coupled to a Q Exactive HF Hybrid Quadrupole-Orbitrap (Thermo Fisher Scientific, Waltham, MA) mass spectrometer. Peptide mixtures were separated by using a 60-min gradient of 92% A (0.1% FA in water) and 8% B (0.1% FA in acetonitrile) to 33% B in 44 min at 300 nl/min with a BEH C18 analytic column (130 Å pore size, 1.7␮m, 75 ␮m⫻ 250 mm; Waters). Peptides were selected for fragmentation automatically by data-dependent analysis.

Protein identification and label-free quantification.The acquired raw files were imported into the Progenesis LC-MS software (v2.0; Nonlinear Dynamics Limited), which was used to extract peptide precursor ion intensities across all samples by applying the default parameters. The proteins were identified by using MASCOT against the predicted proteomes of bothE. coliK-12 (UniProt version July

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2016; narrowed toE. colistrain K-12) andE. coliBL21(DE3) (UniProt version July 2016; narrowed toE. coli strain B/BL21-DE3). The K-12 database consists of 4,303E. coliproteins. The BL21(DE3) database consists of 4,159E. coliproteins. The search criteria were set as follows: fixed modifications, carbamidomethyl (C); variable modifications, oxidation (M); peptide tolerance, 5 ppm; MS/MS tolerance, 0.5 Da; peptide charges, 2⫹and 3⫹. Peptide identification was performed against both theE. coliK-12 proteome and the BL21(DE3) proteome (experimental). Since only a small number of additional proteins were identified, all subsequent analysis was performed with the K-12 reference proteome because of its superior annotation.

Statistical analysis for detection of differentially expressed proteins.Before statistical analysis was performed, QN was applied to the proteomics data sets (69). QN creates equal distribution across all samples by replacing each data point with the mean of the corresponding quantile. It was performed through the R/Bioconductor package preprocessCore (70). PCA of all 15 samples (three conditions, five replicates) was performed to identify condition clustering; one outlier (GI-51) was removed from the data set and subsequent analysis. The presence of differentially expressed proteins was investigated for all possible comparisons (GI-EUI, EI-EUI, and GI-EI) by RS analysis (31) performed with the R/Bioconductor package RankProd (71, 72). ThePvalues obtained were then corrected for multiple testing by the Benjamini-Hochberg method (73).

iGA.The iGA method (32) was used to detect overrepresented (enriched) groups of features for each experimental comparison (GI-EU, EI, EU, or GI-EI). ThePvalues obtained from the iGA were corrected for multiple testing by the Bonferroni method (74). The protein annotation reported in the STEPdb database (34) was used to assign a specific subcellular topology (location) to each protein. iGA was applied to the list of detected proteins ordered according to the one-tailedPvalues obtained from the RS analysis. In accordance with the topology analysis, enrichment analysis of GO terms in the upregulated (or down-regulated) proteins in each comparison was performed by considering all of the GO terms reported in the AmiGO database (33) with two or more members on the list of detected proteins. iGA analysis was used to assess if the detected proteome level changes constitute a unique signature or if they are related to previously studied processes. We collected previously identified transcript/protein signatures for a number of different studies ofE. coli(wild type and/or knockout strains) (Table S3).

Accession number(s).The full data from this study were submitted to ProteomeXchange under project accession no.PXD006474(project doi:10.6019/PXD006474).

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at

https://doi.org/10.1128/mBio

.02118-17

.

FIG S1

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FIG S2

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FIG S3

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FIG S4

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FIG S5

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FIG S6

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FIG S7

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TABLE S1

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TABLE S2

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TABLE S3

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ACKNOWLEDGMENTS

This research was supported by Biotechnology and Biological Sciences Research Council

(BBSRC) grant BB/K014773/1 and BBSRC BioCatNet grant PoC-012 BB/L013649/1. N.D. holds

BBSRC David Phillips fellowship BB/K014773/1. L.G.H. is supported by EPSRC CDT grant

EP/I033270/1. H.M. and R.G. thank the European Commission Seventh Framework

Program for funding (STREPSYNTH; project 613877).

We thank Matthew Bycroft (Reddy’s Laboratories [EU] Ltd.) for providing the AcDKR

clone and activity assay, Mark Blight (Biocatalysts Ltd.) for kindly providing the PL clone,

and the Bio-MS core facility for proteomics analysis.

R.M. planned and performed the experiments, analyzed the data, and wrote the

manuscript. F.D.C. performed the statistical/signature analysis and wrote the

manu-script. H.M. performed the metabolomics analysis and wrote the manumanu-script. L.G.H.

performed experiments. S.H. performed the FACS experiments. R.G. wrote the

manu-script. R.B. and N.D. planned the experiments, analyzed the data, and wrote the

manuscript.

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